More than colocalizing with polycystin-1, polycystin- L is in the centrosome - 《American Journal of Physiology》美国生理学杂志干细胞之家



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发表于 2009-4-22 08:38 |只看该作者 |倒序浏览 |打印

作者：Eva-Flore Bui-Xuan, Qiang Li, Xing-Zhen Chen, Catherine A. Boucher, Richard Sandford, Jing Zhou, and Nuria Basora作者单位：1 Département de Physiologie et Biophysique, Faculté de Médecine et des Sciences de la Santé, Université de Sherbrooke, Sherbrooke, Québec; 2 Membrane Protein Research Group, Department of Physiology, University of Alberta, Edmonton, Alberta, Canada; 3 Depar

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      【摘要】
   Polycystin-1 and polycystin-2 are involved in autosomal dominant polycystic kidney disease by unknown mechanisms. These two proteins are located in primary cilia where they mediate mechanosensation, suggesting a link between cilia function and renal disease. In this study, we sought to characterize the subcellular localization of polycystin- L, a closely related member of polycystin-2, in epithelial renal cell lines. We have shown that endogenous polycystin- L subcellular distribution is different in proliferative and nonproliferative cultures. Polycystin- L is found mostly in the endoplasmic reticulum in subconfluent cell cultures, while in confluent cells it is redistributed to sites of cell-cell contact and to the primary cilium as is polycystin-1. Subcellular fractionation confirmed a common distribution of polycystin- L and polycystin-1 in the fractions corresponding to those containing the plasma membrane of postconfluent cells. Reciprocal coimmunoprecipitation experiments showed that polycystin- L was associated with polycystin-1 in a common complex in both subconfluent and confluent cell cultures. Interestingly, we also identified a novel site for a polycystin member (polycystin- L ) in unciliated cells, the centrosome, which allowed us to reveal an involvement of polycystin- L in cell proliferation.
      【关键词】 cilia cell proliferation autosomal polycystic kidney disease& H) y6 h! f& j0 `4 F& B* Q
   POLYCYSTINS ARE A NOVEL class of transmembrane proteins, whose founding members were polycystin-1 (PC1) and polycystin-2, coded by PKD1 and PKD2, respectively. To date, eight mammalian polycystins have been discovered and can be separated into two categories according to their predicted structure. The first is the PC1-like subfamily which includes PC1 ( 10, 19, 20 ), and the protein products of PKD1L1 ( 57 ), PKD1L2, PKD1L3 ( 25 ), and PKDREJ ( 18 ). Each of these has 11 predicted transmembrane-spanning regions, a large extracellular NH 2 terminus and a short cytoplasmic COOH terminus. The second is the polycystin-2-like subfamily which includes polycystin-2 ( 33 ) and the products of PKDL/PKD2L ( 40, 54 ) and PKD2L2 ( 15, 52 ). Their predicted protein structure, containing six transmembrane-spanning regions with intracellular NH 2 and COOH termini, has such a strong similarity to the transient receptor potential (TRP) channels, that the polycystin-2-like subfamily is now included in the TRP superfamily (TRPP) ( 17, 36, 37 ).
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Mutations in either PKD1 or PKD2 cause autosomal dominant polycystic kidney disease (ADPKD). ADPKD is one of the most common human monogenic disorders (1/400 to 1/1,000) in which the renal parenchyma is progressively replaced by epithelial-lined, fluid-filled cysts. Many other abnormalities are also associated with the disease, including nonrenal cysts (in the liver, pancreas, testes, ovary, and spleen) and cardiovascular and brain complications. Although several distinct biochemical functions have now been associated to either PC1 or -2, their physiological roles remain unknown.

PC1 has been found in various subcellular regions. First, PC1 was located in the plasma membrane in both apical and basolateral compartments. Several groups have reported that PC1 was associated with intercellular junctions at sites of cell-cell contact at either the adherens junction ( 16, 53 ) or in desmosomes ( 40, 44, 46 ). Experiments using Madin-Darby canine kidney (MDCK) cells grown in three-dimensional collagen gels, a cyst/tubulogenesis model, revealed that PC1 was intracellular in MDCK cysts and, as tubules formed, its expression was increased and became located to desmosomes in the plasma membrane ( 40, 45 ). It has been proposed that association of PC1 with desmosomes was important for signaling or for cell adhesion ( 46 ). Some reports have also found PC1 to be colocalized at focal adhesions suggesting a role in mediating cell-matrix interactions ( 12, 53 ). Second, PC1 was located in primary cilia where it modulates intracellular calcium levels in response to fluid flow ( 39 ).
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The precise subcellular distribution of polycystin-2 is still controversial and several lines of biochemical evidence support its residence both in the basolateral plasma membrane and in the endoplasmic reticulum (ER) ( 6, 11, 23, 31, 45 ). Polycystin-2 has also been found in the primary cilia ( 38, 41, 56 ) and, in renal cells, this localization requires PC1 ( 38 ). Accumulating evidence suggests that polycystins-1 and -2 interact to form a channel complex that is involved both in the initiation and maintenance of a terminally differentiated state of tubular epithelial cells, and mechanosensation in relation to fluid flow ( 38 ).

Primary cilia are formed when cells are either in G1 interphase or in quiescent G0. The mother centriole of the centrosome becomes the basal body from which the primary cilium forms and through which ciliary proteins must transit. During the cell cycle, the primary cilium disassembles and the mother and daughter centrioles regroup to form the centrosome. In proliferating cells, centrosomes coordinate the organization of the microtubule bipolar mitotic spindles essential for chromosome segregation. In confluent and nonproliferating cells, the centrosome is responsible for the nucleation of microtubules. Accumulating data have also made it apparent that the centrosome most likely plays an important role in the regulation of the cell cycle ( 1, 9 ).
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Loss of a functional PC1/-2 complex in the primary cilium is believed to be one possible reason for the development of ADPKD. Intense research has focused on PC1 and polycystin-2, while other polycystin members have been largely ignored.

PKDL was the third member of this family to be cloned and, unlike the first two, is not directly linked to ADPKD ( 39, 54 ). PKDL codes for polycystin-Like [polycystin- L ( PCL )], an 805-aa protein that is 50% identical and 71% homologous in amino acid sequence to polycystin-2. PCL functions as a calcium-regulated, calcium-permeable nonselective cation channel when overexpressed in Xenopus laevis oocytes ( 8 ), but its physiological function, as well as that of other polycystins, remains unknown. The expression of PCL is increased in the adult mouse kidney, compared with fetal tissue, where PCL is predominantly found in the apical region of the principal cells of inner medullary collecting ducts ( 2 ).

The aim of the present study was to establish PCL subcellular localization in renal cell lines to extend the current knowledge of polycystin protein function. Using specific antibodies against PCL and PC1, we compared their localization patterns in proliferative and nonproliferative inner medullary collecting duct (IMCD) and MDCK cell cultures. Data presented in this study extend some of the known polycystin characteristics to PCL and revealed a new feature specific for this protein, which allowed us to predict its potential physiological function.& X8 ~8 c  S1 [+ y$ a( f

MATERIALS AND METHODS
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Cell culture. Rat and mouse IMCD cells [rIMCD ( 47 ) and mIMCD-3; American Type Culture Collection (ATCC), Manassas, VA] and MDCK cells (ATCC) were cultured in DMEM supplemented with 10% vol/vol fetal bovine serum (BioWhittaker, Walkersville, MD) and 1% vol/vol glutamine (Cambrex, Walkersville, MD).

Direct and indirect double-labeling immunofluorescence. For immunofluorescence experiments, mIMCD-3 and MDCK confluent cells were plated at high density on glass coverslips and grown for 2-4 days before fixation, and subconfluent cells were plated at 10% confluency and fixed the next day. For all indirect immunofluorescence experiments, except for PC1 and PCL cilia localization, we used the following conditions: mIMCD-3 and MDCK cells were fixed for 10 min with 100% cold methanol (prechilled at -20°C). The cells were washed three times with PBS for 10 min each and blocked with 2% BSA/PBS. The cells were incubated for 1 h with the appropriate primary antibodies or marker: polyclonal anti-PCL [1/100 ( 2 )], polyclonal anti-PC1 (1/500, unpublished PC1 polyclonal antibody against the LRR domain generated by Dr. R. Sandford), monoclonal anti-E-cadherin, clone 36 (1/500, BD Biosciences, Mississauga, ON), monoclonal anti-desmosomal protein, clone ZK-31 (1/250, Sigma, St. Louis, MO), concanavalin A (5 µg/ml, Molecular Probes, Eugene, OR), anti- -tubulin (1/1,000, Sigma), and anti-acetylated -tubulin (1/2,000, Sigma). A commercial rabbit polyclonal anti-PCL antibody (Chemicon, Temecula, CA) was used for mIMCD-3 cilia staining only. The conditions used were those defined by the manufacturer. PC1 staining in the cilia was done in rat IMCD cells, which were seeded onto 0.4-µm polycarbonate membrane transwells (Corning, Acton, MA) and grown for 5 days before fixation in 4% paraformaldehyde and permeablized with 0.1% Triton X-100 (Roche, Laval, QC).

Cells were washed three times for 10 min in PBS and incubated with the appropriate secondary antibodies: anti-rabbit-FITC or -rhodamine conjugated, anti-mouse-FITC or rhodamine-conjugated (Chemicon). After three additional washings, nuclei were stained with DAPI (2 nM, Molecular Probes), and glass coverslips were mounted on glass slides with Vectashield mounting medium (Vector Labs, Burlingame, CA). Subconfluent cells were examined at high magnification on a Nikon Eclipse TE300 microscope (Nikon, Mississauga, ON) equipped with epifluorescence and a stepper motor allowing image acquisition every 0.1 µm. For image acquisition, a CoolSNAP fx digital camera (Roper Scientific, Tucson, AZ) was used. Fluorescence of confluent cells was either observed at x 40 with a Leica DMR/IRBE inverted microscope equipped with epifluorescence (Richmond Hill, ON) and pictures were taken with a Princeton micromax CCD 5-MHz camera, or with the Nikon Eclipse TE300 microscope. PCL staining in cilia was observed at x 100 with the Nikon Eclipse TE300 microscope and PC1 staining in the cilia with a Zeiss LSM510 Meta confocal microscope (Jena, Germany). Pictures were analyzed and merged using MetaMorph software (Universal Imaging, Downingtown, PA) or Simple PCI (Compix, Imaging Systems, Cranberry Township, PA).8 Z! D. N1 q: X/ j% i7 h2 l
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Immunoprecipitation and coimmunoprecipitation. Cells from a 100-mm petri dish (80-90% subconfluent or 4 days postconfluent) were lysed for 1 h in 1 ml of cold lysis buffer containing 1% Triton X-100, 50 mM Tris, 150 mM NaCl, and Complete mini EDTA-free protease inhibitors (Roche) and assiduously vortexed every 15 min. Insoluble material was removed by centrifugation at 13,000 rpm for 15 min at 4°C. Lysates containing 4 mg of protein in 1 ml were precleared by incubation with protein G agarose (Roche) for 1 h at 4°C on a rotator. Cleared supernatants were incubated overnight with polyclonal anti-PCL (1/500) or anti-PC1 (1/500) sera or with protein G beads alone. Protein G agarose was then added and lysates were incubated for an additional hour. Immunoprecipitates were washed once with lysis buffer and twice with buffer containing 50 mM Tris/150 mM NaCl, eluted with 25 µl solubilization buffer, and heated for 10 min at 100°C. Immunoprecipitates (50% of the sample) and lysates (0.03% of the Triton soluble lysate) were separated on a 7% SDS-PAGE gel, transferred onto nitrocellulose (Hybond ECL, Amersham-Pharmacia, Baie-Urfe, QC), blocked with 5% milk powder/PBS, and probed with polyclonal anti-PCL (1/500) or anti-PC1 (1/500) sera followed by a horseradish peroxidase-conjugated anti-rabbit secondary antibody (Amersham) and detected by chemiluminescence (ECL, Amersham). Protein size was compared using Kaleidoscope prestained protein standards (Bio-Rad, Hercules, CA).

Biotinylation of cell-surface proteins of subconfluent mIMCD-3 cells. Subconfluent mIMCD-3 cells were grown in 100-mm dishes. The cells were placed on ice and washed twice with ice-cold extracellular medium (140 mM NaCl, 1.8 mM CaCl 2, 1 mM MgCl 2, 15 mM HEPES, pH 7.4) and then incubated with gentle rotation for 30 min at 4°C with 2 mg/ml Sulfo-NHS-SS-Biotin (Pierce-BioLynx, Brockville, ON). The biotinylation reaction was terminated by washing the cells three times with ice-cold PBS (137 mM NaCl, 3.5 mM KCl, 0.9 mM CaCl 2, 1 mM MgCl 2, 10 mM Na 2 PO 4, pH 7.4) containing 10 mM glycine. The cells were then lysed with 1 ml of ice-cold lysis buffer (1% Triton X-100, 50 mM Tris, 150 mM NaCl, and Complete mini EDTA-free protease) for 30 min at 4°C followed by 20 passages through a 20-gauge needle and 15 passages through a 25-gauge needle. The cell extracts were cleared by centrifugation and added to 100 µl of streptavidin-agarose beads (Pierce) and incubated for 16 h at 4°C. The biotin-streptavidin-agarose complexes were harvested by centrifugation and washed once with ice-cold lysis buffer and twice with ice-cold washing solution (50 mM Tris, 150 mM NaCl). The beads were then resuspended in 2 x solubilization buffer and incubated at 60°C for 30 min before SDS-PAGE fractionation and Western blotting. One percent of the Triton soluble fraction was loaded for quantification of total protein amount and approximately one-half of the biotinylated sample was loaded to quantify the proportion of membrane protein.
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Subcellular fractionation of confluent mIMCD-3 cells. Cells from two 100-mm petri dishes (10 days postconfluent) were rinsed three times with PBS and once with the wash buffer (0.25 M sucrose, 10 mM triethanolamine-acetic acid, pH 7.8). Cells were scraped into 1 ml of homogenization buffer (0.25 M sucrose, 10 mM triethanolamine-acetic acid, pH 7.8, 1 mM EDTA) and homogenized using 23 pestle strokes of a 3 ml Dounce homogenizer. The 0-26% (wt/vol) 10 ml linear iodixanol gradient (Optiprep, Axis-shield, Oslo, Norway) was prepared using a two-chamber gradient maker (low-density end first) and refrigerated 1 h before use. The sample was loaded onto the top of the gradient and spun for 120 min at 40,000 rpm at 4°C in a SW.41 rotor in a Beckman ultracentrifuge. Fractions ( 500 µl each) were collected from the bottom using a peristaltic pump with a dropwise fraction collector. Equal volumes of each fraction were subjected to SDS-PAGE on 10% Laemlli gels followed by immunoblotting analyses. Antibodies used for fraction analyses were anti-calnexin for the ER (clone 37, 1/1,000, BD Biosciences), anti-E-cadherin (1/2,000) for the plasma membrane, anti-PCL (1/500, Chemicon), and anti-PC1 (1/500).

Study of in situ subconfluent cell proliferation. mIMCD-3 cells were plated on glass coverslips the day before transfection as described above. Cells were transfected following the manufacturer?s protocol for Lipofectamine 2000 (Invitrogen Life Technologies, Carlsbad, CA) with a human PKDL construct containing an in-frame NH 2 terminal, Xpress tag in pcDNA3.1HisC (PCLX), and an HA-tagged calreticulin construct ( 14 ) as an overexpression control. Twenty-four-hour posttransfection cells were loaded with 10 µM BrdU (In situ Cell Proliferation Kit, fluos; Roche) for 2 h and subsequently treated as described ( 35 ). Transfected cells were visualized by indirect immunofluorescence using anti-Xpress (1/500, Invitrogen) or anti-HA (clone 12CA5, 1/250, Roche) and BrdU-positive cells were visualized using anti-BrdU (1/5, Roche). Ten to 12 fields for each condition in 3 independent experiments were manually analyzed. Data were compiled and subjected to statistics using the z -test to compare the two groups for significant difference of proportion (SigmaStat, SPSS).
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Expression and distribution of PCL. Characterization of the anti-PCL serum and the expression and distribution patterns of PCL in the developing and adult mouse kidney have been reported ( 2 ). In an attempt to further characterize the subcellular distribution of endogenous PCL in this context, we used different kidney epithelial cell lines: IMCD and MDCK cells. Results for both cell lines were identical and, for simplification, only results for IMCD cells are shown.

We compared, in parallel, the localization of PC1 and PCL with that of various cell structure markers: E-cadherin, an adherens junction marker, a desmosomal protein recognized by monoclonal ZK-31, acetylated -tubulin, which labels primary cilia and concanavalin A, a lectin capable of binding sugar groups normally found in the ER.7 b' `# b3 J% }
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We verified the validity of two chosen markers in our study with a double-labeling immunofluorescence experiment in subconfluent IMCD cells ( Fig. 1 A ). As expected, E-cadherin was in the membrane at sites of cell-cell contact in subconfluent cell cultures ( Fig. 1 A, green panel) and there was no overlapping with the concanavalin A staining ( Fig. 1 A, red panel and merged image).6 I' L- i( m8 K
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Fig. 1. Localization of polycystin-1 (PC1) and polycystin- L (PCL) in subconfluent cells. Localization of PC1 and PCL was determined by indirect immunofluorescence microscopy in mIMCD-3 cells using immunosera raised against PC1 and PCL. Controls ( A ) confirmed that no overlap occurred between the plasma membrane marker E-cadherin ( A, green panel) and the endoplasmic reticulum (ER) marker concanavalin A (conA; A, red panel) on subconfluent mIMCD-3 culture. In mIMCD-3 cells, PC1 ( B, top green panel) was mostly intracellular and costained with conA ( B, bottom red panel) in the ER and was weakly detected in the plasma membrane ( B, bottom ). PCL staining ( C, green panels) also overlapped with conA ( C, top ) in the ER and a faint plasma membrane staining was also seen ( C, bottom ). All pictures were taken at x 100.

Expression and localization of PCL were compared with that of PC1 in IMCD subconfluent cultures ( Fig. 1, B and C ). In IMCD cells, endogenous PC1 staining ( Fig. 1 B, top green panel) appeared to be mostly intracellular and codistributed with concanavalin A ( Fig. 1 B, top red panel). Comparison with E-cadherin showed that PC1 ( Fig. 1 B, bottom green panel) was weakly present at the plasma membrane at sites of cell-cell contact where intercellular junctions have begun to form. Endogenous PCL ( Fig. 1 C, top green panel) showed a similar intracellular staining pattern with expression codistributing extensively to the ER with concanavalin A ( Fig. 1 C, top red panel) and was also visible in the plasma membrane, evidenced by codistribution with E-cadherin ( Fig. 1 C, bottom green panel). Interestingly, one or two bright dots were invariably seen with the anti-PCL serum at a different plane of focus than that observed for E-cadherin, and they were suspected to be associated with centrosomes or centrosomal structures. We did not observe, a characteristic focal adhesion staining pattern for either protein, in our cell models.

As expected for confluent cell cultures, PC1 colocalized with acetylated -tubulin in primary cilia ( Fig. 2 A, a-c ). PC1 redistributed to the basolateral plasma membrane and partially overlapped with both E-cadherin ( Fig. 2 A, d-f ) and the desmosomal protein ZK-31 ( Fig. 2 A, g-i ), but was not readily detectable in the ER ( Fig. 2 A, j-l ). Careful analyses of 0.3-µm z -section images revealed that PC1 distribution overlapped more closely at planes of focus of desmosome. Interestingly, PCL staining partially overlapped with PC1 distribution in confluent cell cultures. PCL was indeed found in primary cilia as showed by the colocalization with acetylated -tubulin ( Fig. 2 B, a-c ). Labeling was punctuated along the entire length and was also clearly visible in the basal body and daughter centriole at the base of the cilia. At lower focal planes, PCL was detected in the basolateral plasma membrane as determined by the codistribution with E-cadherin ( Fig. 2 B, d-f ) and with the desmosomal protein ZK-31 ( Fig. 2 B, g-i ). Acquisition of 0.3-µm stacked images and careful analysis of these images revealed that PCL plasma membrane staining overlapped more prominently with desmosomal junction than with E-cadherin. A subpopulation of PCL was also present in the ER, as demonstrated by the costaining with concanavalin A ( Fig. 2 B, j-l ).& u" l. v2 O+ E+ `' x
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Fig. 2. Common distribution of PCL and PC1 in confluent inner medullary collecting duct (IMCD) cells. Localization of PC1 and PCL was determined by immunofluorescence microscopy in polarized IMCD cells at 4 days postconfluence, using a PC1 ( A, a, d, g, j ) or a PCL ( B, a, d, g, j ) antiserum. As expected, PC1 was located to the cilia as seen by colocalization with acetylated -tubulin (tub; A, a-c ). The majority of PC1 was found in the basolateral plasma membrane with the adherens junctions marker, E-cadherin (Ecad; A, d-f ), and with a desmosomal protein (ZK31; A, g-i ). PC1 displayed a punctuate staining along the membrane, a characteristic associated with desmosomes. No intracellular staining overlapping with concanavalin A (conA; A, j-l ) was visible. PCL distribution was highly similar to PC1 and was found in the cilia as seen by colocalization with acetylated -tubulin (tub; B, a-c ). PCL was found in the basolateral plasma membrane with the adherens junction marker Ecad ( B, d-f ), and with a desmosomal protein (ZK-31; A, g-i ). PCL could be seen in discrete, punctuate staining along the membrane. A larger proportion of PCL was seen as intracellular staining overlapping with conA ( B, j-l ). Images were taken at different focal planes for the different cell structure markers. All pictures were taken at an original magnification of x 40, except desmosome pictures and cilia pictures ( x 100): PC1 cilia images were acquired with a confocal, while PCL cilia images were acquired at x 100 with a microscope epifluorescence equipped.6 G, W! E! ]5 J( _4 d" \9 P- [! V8 L
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To measure the proportion of the amount of PCL residing in the plasma membrane of subconfluent IMCD cells, which was weakly visible by indirect immunofluorescence, we carried out biotinylation of the cell surface in subconfluent cell cultures ( Fig. 3 A ). PCL was detected in the biotinylated fraction, albeit in relatively small amounts ( Fig. 3 A, right ). We calculated the proportion of PCL at the cell surface of subconfluent cells to be 0.5% of the total amount. This was determined by estimating band intensity in relation to the proportion of protein loaded. This confirmed the immunostaining, which showed that the large majority of PCL was intracellular in subconfluent cells. The integrin 1-subunit, a well-characterized resident plasma membrane protein used as a positive control, was greatly enriched in the biotinylated fraction ( Fig. 3 A, left ).( j! U/ r) w4 a0 T) v6 J
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Fig. 3. PCL is found in the plasma membrane in proliferative cells and forms a complex with PC1 in confluent mIMCD-3 cells. A : cells at 90% confluency were labeled with NHS-Biotin and solubilized in 1% Triton. Fifty micrograms of protein from the Triton-soluble fraction (Ly) and one-half of the total protein from the streptavidin purified fraction (Biotin) were detected for the integrin 1 subunit or PCL. B : confluent cell lysates (4 mg/ml) were immunoprecipitated (IP) using anti-PC1 serum (PC1; 1/500), anti-PCL serum (PCL; 1/500), and were subsequently immunoblotted (IB) with anti-PC1 serum ( top ) or anti-PCL serum ( bottom ). NS, nonspecific control, lysate with only protein G beads. PC1 migrated at 500 kDa; PCL migrated as 2 forms, which migrate at 90 and 120 kDa. The two bands were present in this experiment because the amount of sample loaded allows for detection of the weaker 120 kDa species. The nature of this band is not known but is detected with two different antibodies to PCL raised against different epitopes. C : subcellular fractionation of confluent IMCD cells using a 0-26% iodixanol gradient. PCL co-sedimented with the ER marker calnexin in the dense fractions ( bottom ) and with the plasma membrane marker E-cadherin in the lighter fractions. PC1 showed a broader distribution and overlapped fractions containing E-cadherin, but not calnexin./ _% i+ E6 \  d
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We used reciprocal coimmunoprecipitation studies to determine whether PC1 and PCL could be found in a common complex in confluent cell cultures ( Fig. 3 B ). Anti-PC1 detects a band migrating at 500 kDa in both the soluble lysate fraction and the immunoprecipitate using the anti-PC1 serum ( Fig. 3 B ). This band was also detected with anti-PC1 following immunoprecipitation using the PCL antiserum, confirming that these two endogenous proteins are found in a common complex. Native PCL migrated as two distinct bands with molecular weights at 90 and 120 kDa and may represent alternatively spliced forms ( 13, 28 ). Identical bands were obtained using recombinant PCL overexpressed in HEK cells (data not shown). Reciprocal coimmunoprecipitation of PCL was observed when using the anti-PC1 sera followed by detection with the PCL serum ( Fig. 3 B ). Identical results were obtained using MDCK cells and in subconfluent cell cultures of both cell lines (data not shown).
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To extend the coimmunoprecipitation results and to confirm the immunofluorescence carried out with a methanol fixation, that could create artefacts, we undertook subcellular fractionation on 10-day postconfluent IMCD cell cultures. Cell lysates were separated on a 0-26% iodixanol gradient and fraction samples were analyzed by SDS-PAGE ( Fig. 3 C ). Anti-calnexin, a resident ER protein used to identify which fractions contained the ER membranes, was found in the dense fractions at the bottom of the gradient. Plasma membrane marker E-cadherin could be found in the top two-thirds of the gradient corresponding to the lighter fractions.7 P/ L" q9 x% D. L- j6 Y+ r

A minor portion of PCL was detected in the densest fractions containing the highest concentration of ER membranes. The majority of PCL was found in the lighter fractions comigrating with E-cadherin and could also be seen at higher molecular weights, possibly representing dimers and multimers.

PC1 was detected in the lighter fractions overlapping with E-cadherin. Its distribution in these fractions appeared broader than that observed for PCL. PC1 was not, however, detected in fractions containing the ER membranes. A faster migrating species was also observed with anti-PC1 and most likely represents previously described cleavage products ( 7, 42 ). Thus PCL and PC1 were found together in the lightest fractions of the plasma membrane, which supports the coimmunoprecipitation results, and PCL was found alone in the ER, confirming the data obtained by indirect immunofluorescence.

We next sought to further characterize suspected colocalization of PCL with the centrosome, first observed in the subconfluent cultures of both cell lines studied. We used anti- -tubulin as a centrosome marker and DAPI to identify the different phases of the cell cycle ( Fig. 4 ). Anti-PCL serum labels a bright dot located in the perinuclear region of cells in interphase. This bright dot costains perfectly with anti- -tubulin, confirming centrosome association of PCL. As the cell enters prophase, PCL is clearly seen in the duplicated centrosomes, which are now visible as two bright dots and have begun to move apart over the nucleus.4 Y: N% |  D, O1 N) s# U* ~! n

Fig. 4. PCL localizes to the centrosomes. Proliferating cell cultures of mIMCD3 cells were subjected to indirect immunofluorescence staining using anti-PCL serum (green) and anti- -tubulin (red). Nuclei were stained with DAPI and used to identify the stage of cell cycle division. Inset : for those images where the centrosomes were not in the same plane of focus, the second centrosome is shown. Arrows mark the location of these centrosomes. All images were taken at x 100.$ {# i# w: k8 l  e) L9 L, i5 Z6 Z
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The centrosomes are positioned on either side and remain positively labeled for PCL from prometaphase through anaphase. Insets show, when necessary, the second centrosome located in a different plane of focus than the first. Telophase marks the end of the cell cycle and results in the generation of two daughter cells each with one centrosome with which PCL is still clearly labeled. Therefore, throughout all stages of the cell cycle, PCL was colocalized with -tubulin in the centrosome. Moreover, levels of PCL associated with the centrosomes appeared to remain constant as division occurs, suggesting that this location is an intrinsic characteristic of PCL.) l; |' x9 n( q

A recent report revealed that polycystin-2 does not associate with the centrosome in cycling cells (i.e., unciliated cells), but rather, was associated with mitotic spindles ( 43 ). We carried out double staining immunofluorescence experiments in proliferating IMCD cells using anti-acetylated -tubulin and anti-PCL serum.

During interphase, anti-acetylated- -tubulin labels a portion of the microtubule cytoskeleton (poorly visible at this plane of focus) and a portion which remains associated around the centrosome. PCL was located, as reported above, to a bright puncta found in the perinuclear region. At prophase, two punctuate regions which are positively stained with both antibodies indicate that centrosomes are duplicated. The interphase microtubule network is disassembled and acetylated -tubulin accumulates more intensely at the centrosome. Once in prometaphase, acetylated -tubulin was not seen at the centrosome but was found in the newly forming bipolar mitotic spindles. PCL was clearly visible in the centrosome at the origin of mitotic spindles but did not appear to codistribute with stabilized microtubules at this stage. This pattern of distribution was also observed at metaphase when the microtubule spindles are seen reaching into the aligned chromosomes and as the chromosomes move apart in anaphase. During telophase and cytokinesis, anti-acetylated -tubulin staining appeared as weak puncta and these structures were positive for PCL. Intense staining with anti-acetylated- -tubulin was seen in the midbody (telophase inset) but still no PCL was detectable with these microtubules ( Fig. 5 ).0 j& M4 a5 B* \# Y

Fig. 5. PCL does not colocalize with the mitotic spindle. Proliferating cell cultures of mIMCD3 cells were subjected to indirect immunofluorescence staining using anti-PCL serum (green) and anti- -tubulin (red). Nuclei were stained with DAPI and used to identify the stage of cell cycle division. Inset : for those images where the centrosomes and the mitotic apparatus were not in the same plane of focus, the second centrosome are shown. Arrows mark the location of those centrosomes found in a different focal plane. All images were taken at x 100." j* T( j+ P5 f
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Proteins associated with the centrosomes are quite often involved with the regulation of cell growth. Overexpression of both PC1 and polycystin-2 has been shown to inhibit cell growth ( 4, 30 ) and we questioned whether PCL may also share this characteristic. We generated a tagged human PDKL sequence carrying an Xpress tag in the NH 2 -terminal domain. We used BrdU incorporation assays to determine whether cell proliferation was affected by overexpression of tagged PCL in transient transfection conditions. Transfected cell populations were identified using the anti-Xpress tag 24 h after transfection in cells that had been labeled with BrdU ( Fig. 6 A ). We used the ER resident protein calreticulin-HA as a negative control for overexpression effects ( Fig. 6 A ) ( 32 ). Experiments showed that 39.4% of calreticulin-transfected cells were proliferating ( Fig. 6 B ). Transient transfection of PCL resulted in an important decrease of cells in S-phase (16.5%) and this 2.4-fold decrease was shown to be statistically significant ( P " f: `1 t! W' p

Fig. 6. Overexpression of PCL reduces cell proliferation. A : transient transfection of IMCD cells with calreticulin-HA ( a ) or PCLX ( b ) labeled with BrdU. Transfected cells (red) were visualized using anti-HA (calreticulin) or anti-Xpress (PCLX) while proliferative cells were identified with anti-BrdU-FITC (green). All nuclei were stained with DAPI (blue). Representative images for each condition are shown. B : cells in a minimum of 10 fields were counted to determine the total number of transfected cells (red) and the number of BrdU-labeled transfected cells (red and green) from 3 independent experiments. Results are presented as % of proliferative transfected cells = # of red and green cells/total # of red cells * 100. In all, between 1,300 and 1,600 cells were counted for each condition and statistics ( z -test) were done so as to compare proportions between 2 groups for significant difference (* P

DISCUSSION& F' R' v4 ~& A
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In this study, we showed that PCL distribution largely overlapped with, and was associated to PC1, a known human disease gene. PCL was found in the plasma membrane and in the primary cilia of renal epithelial cells. Coimmunoprecipitation experiments suggested that in these common areas of PCL and PC1 expression they reside in the same protein complex. A novel finding is the presence of a polycystin member, PCL, in the centrosomes of proliferative and in confluent, nonproliferative cells.0 C- f+ Y- c2 `# Q. }

Localization of PC1 was somewhat controversial and has been reported in several subcellular compartments (for a review, see Ref. 5 ), and in fact, the same is true for polycystin-2, whose localization is also currently under debate (for a recent review, see Ref. 22 ). Our study showed that both PC1 and PCL were identified in the primary cilia, confirming previous studies on PC1 and suggesting that residence in cilia is a characteristic of polycystin family members. Distribution of PC1 was more extensive with the desmosomes than with E-cadherin. In fact, close inspection of PC1 staining in the cell lines studied here revealed a "beads on a string" pattern, characteristic of desmosomal components and has been shown by others ( 46 ). PCL also displayed a highly similar punctuate pattern, like the desmosomal structures. A significant fraction of PCL was also found in the ER. Whether PCL is functional in each of these compartments, as has been suggested for polycystin-2, remains to be determined.

Proper function of PCL, like polycystin-2, may require distinct contexts, possibly in different subcellular compartments. The cytoskeleton is most likely important for polycystin function as experiments aimed at identifying interacting proteins have found a number of actin-binding proteins, which bind to PC1 ( 12 ), polycystin-2 ( 24, 26 ), and PCL ( 27 ) and intermediate filament proteins, which bind to PC1 ( 55 ). This hypothesis is strongly supported by a number of recent papers which report the modulation of polycystin-2 or PCL channel function by actin cytoskeletal components ( 26, 29, 34 ).! T7 h& T! {3 U/ H9 T4 g2 S  ?& P
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Association of PCL with PC1 in a common complex was confirmed by reciprocal coimmunoprecipitation, in both subconfluent and confluent cell cultures. Combined analyses from indirect immunofluorescence and subcellular fractionation experiments suggest that a PC1/PCL complex would reside in the plasma membrane and/or in the cilia. However, the nature of this interaction, whether direct or indirect, remains to be confirmed. In vitro yeast two-hybrid analyses using the PC1 terminus with the PCL terminus failed to identify sites of direct interaction (Q. Li and X. Z. Chen, unpublished observations). It is possible, however, that other regions of PC1 or PCL are required, such as intracellular loops or the transmembrane segments as has been shown for polycystin-2 and TRPC1 interaction ( 49 ).

The putative existence of a PC1/PCL complex is supported by the lack of perfect coexpression between PC1 and its partner, polycystin-2, in various tissues. This led to the belief that, either polycystins could function alone, or that they could associate with other, as yet unidentified proteins, including other members of the polycystin family. It will be important to determine whether polycystin complexes coexist in the same cell types or are cell specific. Attempts to coimmunoprecipitate polycystin-2 and PCL in IMCD cells were unsuccessful (E. F. Bui-Xuan and N. Basora, unpublished data) raising the possibility that PC1 forms distinct complexes with each polycystin-2 and PCL.6 B6 r# [' `7 c/ X: t- a5 r: H& B
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These observations raise the question of what function a PC1/PCL channel complex would have in the plasma membrane at sites of intercellular contact. One hypothesis could be that PC1 monitors desmosome integrity and a polycystin channel complex is activated when this structure is compromised. A membrane-anchored form of the PC1 C-terminal, thought to act as dominant-positive, not only enhances ion channel activity ( 50, 51 ) but can also, as mentioned above, modulate various signaling pathways including PKC. This is especially relevant in light of the fact that desmosomes are sensitive to PKC ( 48 ) and intracellular calcium ( 21 ).

The PC1/2 complex has been shown to modulate intracellular calcium levels in response to fluid flow due to its location in the primary cilia ( 38 ). The impact of the coexistence of a PC1/PCL channel in the same structure is not known. One hypothesis could be that these different channels are activated in response to different rates of flow. This would be analogous to structurally related TRPV vanilloid receptors. TRPV1 and TRPV2 channels are both activated by heat, but at different threshold temperatures; TRPV1 (VR-1) is activated at temperatures greater than 43°C while TRPV2 (VRL-1), which is 50% identical to TRPV1, is activated at temperatures greater that 53°C ( 3 ).$ Q% i2 _4 ?0 I) W0 Q% U

One unexpected novel result to come out of this study is the resident site of PCL to the centrosome in dividing (subconfluent) and nondividing (confluent) cells. It is believed that there are 200 centrosomal or centrosomal-associated proteins, not all of which have been identified. Although centrosomes have long been known to act as microtubule organizing structures, accumulating studies show that centrosomes, via the proteins which associate with them, are most likely involved in various cell processes such as active participation in the coordination of cell cycle initiation/progression ( 9 ), regulation of gene transcription, and protein recycling (ubiquination pathway) ( 1 ). A recent study showed that polycystin-2 was associated with mitotic spindles, but did not appear to be in centrosomes in dividing cells ( 43 ). The exact opposite appears to be the case for PCL, which was undetectable in microtubule structures. The localization of channel proteins to nonmembranous regions of the cell is perplexing but may provide an unsuspected link between polycystins, cell cycle, and epithelial polarity. A possible role for PCL in cell proliferation is underscored by the BrdU incorporation assays. Our results showed that overexpression of an NH 2 -terminal tagged PKDL construct inhibited cell growth, reflected by the significantly reduced number of cells in S-phase. A more extensive characterization is necessary to determine by what mechanism exogenous PCL interferes with the cell cycle and will be the focus of future experiments.% Q7 w3 f+ s5 S$ ]) ]3 K

The results presented in this study on PCL subcellular localization and function highlight the common characteristics between the different family members characterized to date, such as plasma membrane, ER, and cilia localization. Moreover, overexpression studies showed that PCL inhibited cell proliferation. Identifying the similarities and differences between the various family members will undoubtedly offer new insight into the physiological functions of polycystins in general, which are essential in leading to a better understanding of ADPKD.5 [! j2 I( B% U# j

GRANTS
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N. Basora is a recipient of a Fonds de la Recherche en Santé du Québec New Investigator Scholarship award. X.-Z. Chen is a Canadian Institutes for Health Research New Investigator and Alberta Heritage Foundation for Medical Research Scholar.
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ACKNOWLEDGMENTS8 N* P+ n  n/ ~/ F' O

We thank P. Pothier and P. Kilian for assistance with image acquisition. We are also grateful to Troy Sturtevant and Rory Campbell (Bishop?s University Honour?s Program; Lennoxville, Quebec) for technical assistance.
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